Introduction

One of the main challenges in anti-cancer photodynamic therapy (PDT) is to develop novel photosensitizers (PSs) that can preferentially accumulate in malignant tissues relative to normal tissues (1). To achieve this, various approaches have been explored that include: encapsulation in colloidal carriers such as liposomes (2,3) or polymeric micelles (4,5), and conjugation to antibodies (6,7), synthetic peptides (8–10), nanoparticles (11,12) and carbohydrates (1,13–17).

There has been increased interest in the synthesis and development of glycoconjugated PSs because most cancer cells are characterized by increased glucose uptake and overexpress glucose transporters (GLUTs), so as to accommodate the high rates of glycolysis (18,19). Hence, glycoconjugation of the PSs should improve their membrane interaction and uptake by cancer cells, thereby increasing the tumor selectivity of subsequent PDT (1,14,20). In addition, the conjugation of PSs to carbohydrates aids their water solubility, an important parameter for cellular uptake of the PSs (1,21,22).

In this study, we synthesized and chemically characterized zinc phthalocyanines (ZnPcs) conjugated at the macrocycle periphery by four glucose substituents. We examined the uptake kinetics, subcellular localization, in vitro phototoxicity and reactive oxygen species (ROS) generation of our new compounds in MCF-7 cancer cells, compared with disulfonated aluminum phthalocyanine (AlPcS2), a second generation PS currently under clinical evaluation (23).

Synthesis of protected tetra-β-glycosylated ZnPc {4}. A mixture of 4-glycosylated phthalonitrile {3} (5.36 g, 1.6 mmol) and Zn(OAc)2·2H2O (5.14 g, 0.02 mol) in n-pentanol (50 mL) was heated to 100°C, and then DBU (10 mL) was added. The mixture was heated further to 120°C and kept stirring at this temperature overnight. The solvents were then evaporated under vacuum and the residue was purified by silica-gel chromatography using ethyl acetate–n-hexane (changing gradually from v/v 1:3 to 1:1) as the eluent. The product {4} was isolated as a blue–green solid (2.19 g, 85%).

Synthesis of tetra-β-glycosylated ZnPc {5}. Tetra-β-glycosylated ZnPc {5} was obtained by removing the isopropylidene methyl-protecting groups from {4}. To achieve this, compound {4} (1 g, 0.621 mmol) was dissolved in 20 mL ethylene glycol, and then 100 μL trifluoroacetic acid (TFA) was added and the mixture stirred at 120°C overnight. The solvents were evaporated under vacuum and the residue was extensively washed with ether to yield {5}, as a bluish green solid (0.68 g, 85%).

PS loading and PDT treatment. All manipulations prior to PDT were carried out under subdued lighting. The medium containing serum was replaced with 500 μL serum-free medium containing the PS and cells were incubated in the dark at 37°C and 5% CO2 for up to 4 h in initial pharmacokinetic tests, and for 2 h in PDT cytotoxicity studies. The PS-containing medium was then removed and the cells rinsed three times with warm PBS before phenol red-free DMEM supplemented with 1% pen/strep and 10% FCS was added. Test samples were immediately exposed (for 15 min) to 28.6 J cm−2 halogen white light from a 500 W bulb, heat filtered through 5 cm of water in a T175 flask (Nunc 156502) (or maintained in the dark, for dark cytotoxicity experiments). Samples were then returned to the cell culture incubator for a further 24 h post PDT in the dark, and then assayed for viability.

PS uptake. The uptake of the PSs in the total cell population was determined primarily by fluorescence spectroscopy as described previously (24,25). Following incubation with PS for 15 min to 4 h, as detailed above, cells were washed three times with PBS, detached by adding 100 μL of 0.25% EDTA–trypsin per well and the cell number determined using a hemocytometer. Thereafter, 100 μL of 10% sodium dodecyl sulfate (SDS) (to achieve a final concentration 5% v/v) was added to disrupt the cell membranes and samples were briefly vortexed. The samples were gently rocked for 1 h under subdued light conditions, centrifuged at 9200 g for 30 min and the concentration of PS in the supernatant was assayed by measuring the fluorescence using a FLUOstar Optima plate reader (λex = 355 nm, λem = 680 nm). The concentration of PS was determined by comparison with a calibration curve obtained with standard solutions of the same PS in 5% SDS. All experiments were performed in triplicate and were repeated three independent times. The fluorescence values obtained from each sample were corrected to the total number of cells as determined by hemocytometer to allow calculation of the absolute PS concentration per cell. Every experiment was also compared with a culture control without PS (untreated), to determine the level of background fluorescence.

To corroborate the fluorescence spectroscopy PS uptake data by independent means, we used inductively coupled plasma mass spectrometry (ICP-MS; Agilent 7500a) to assay aluminum and zinc ions in cell lysates. About 0.1 mL aliquots of cell lysate from 4 h incubation samples (as detailed above) were mixed with 0.9 mL 2% HNO3 in 5% SDS and used for simultaneous measurement of zinc and aluminum. The instrument is fitted with a standard quartz spray chamber and a PFA nebulizer. Samples were aspirated at approximately 400 μL min−1 with count rates of the order of 3 − 5 × 107 cps ppm−1. Oxide interferences were kept low at 0.5% CeO+/Ce+ (monitored in the tuning solution) and doubly charged species at ≤1% (Ce++/Ce+). Analyses were standardized against a suite of synthetic reference materials, made up from certified plasma standard solutions (Alfa Aesar). The synthetic reference materials were doped with an appropriate aliquot of control cells without PS, in order to minimize the differences in ionization potential between synthetic standards and the natural samples. The detection limit for aluminum was 9 ppb and for zinc was 5 ppb.

Resonance light-scattering measurements. The main theoretical aspects of resonance light-scattering (RLS) method are disclosed in Pasternack and Collings (26). RLS is widely used for the sensitive detection of chromophore aggregates, including PDT dyes (e.g. m-THPC) (27,28).

RLS measurements were performed using a Shimadzu RF-1501 spectrofluorimeter by simultaneous scanning of PS samples with both the excitation and emission monochromators tuned to the same wavelength. Then the recorded RLS spectra were corrected for internal light filter effect and for the sensitivity of the spectrofluorimeter (27). A 5 μm aqueous solution of merocyanine 540 (MC540, Sigma) was used as an aggregation positive control (27,29).

Cell viability analysis. Cells were washed three times with PBS and the collected culture medium and washes were combined to ensure that any detached cells were not lost. The remaining attached cells were removed with trypsin–EDTA and the cell suspension combined with the cells already collected and the total cell number was determined by hemocytometer. The cells were incubated with 20 μg mL−1 propidium iodide (PI) on ice and analyzed by flow cytometry using a FACSCalibur™ cytometer to establish the number of dead (PI positive) cells. For each sample, 10 000 events were acquired on a logarithmic scale and the gating of single cells was achieved by the analysis of forward and side-scatter dot plots using BD CellQuest™ Pro software. PI fluorescence intensity was measured in FL-3 with an emission wavelength of 670 nm. In some experiments, 24 h after illumination, PI was added to live cells to 20 μg mL−1 and photographed with brightfield and TRITC fluorescence on a Nikon SMZ1000 epi-fluorescence stereomicroscope.

Measurement of ROS. ROS levels were determined by flow cytometry using 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA). Cells were seeded in 24-well tissue culture plates at a density of 2 × 105 cells mL−1 and incubated at 37°C overnight and then incubated with 9 μm ZnPc {4}, {5} or AlPcS2 for 2 h. After incubation, the PS-containing medium was removed and the cells rinsed three times with warm PBS. Fresh phenol red-free culture medium with 10 μm DCFH-DA was added under subdued light conditions and the test samples then exposed to 28.6 J cm−2 white light (or not in the case of dark cytotoxicity). The cells were then washed twice with cold PBS, trypsinized and centrifuged for 5 min at 550 g and at 4°C. The cell pellet was resuspended in 200 μL cold PBS and probe fluorescence was measured using FACSCalibur™ cytometer by collecting 10 000 events for each sample. ROS levels were expressed as mean fluorescence intensity as calculated by BD CellQuest™ Pro software.

Results

Synthesis and characterization of glycosylated ZnPcs

The overall aim was to synthesize tetra-β-glycosylated ZnPcs from glycosylated phthalonitriles, as shown in Scheme 1.

The synthesis of 4-glycosylated phthalonitrile {3} was achieved by converting d-glucose into a diacetonide, 1,2:3,4-di-O-isopropylidene-α-d-glucopyranose {1} by protecting the hydroxyl groups via the well-established method of isopropylidene derivative formation (30) and was attached to the phthalonitrile via its position 6. The conventional procedure of protection using the isopropylidene group involves the reaction of the sugar with acetone or with 2,2-dimethoxypropane, both in the presence of a Lewis or Brønsted acid catalyst under anhydrous conditions (31) and in this synthesis, acetone and concentrated sulphuric acid were used. 1H NMR spectroscopy was used to characterize {1} and it gave the characteristic chemical shifts with the glucose CH protons resonating as six distinct multiplets at 3.9–06.0 ppm and the protecting isopropylidene methyl groups resonating as four singlets at 1.2–1.5 ppm (Fig. S1). Thereafter, 4-(1,2:3,4-di-O-isopropylidene-α-d-glucopyranose)-phthalonitrile {3} was prepared from 1,2:3,4-di-O-isopropylidene-α-d-glucopyranose {1} and 4-nitrophthalonitrile {2}, by nucleophilic displacement of the nitro group at 120°C, forming a glycosidic bond, as shown in step 1 of Scheme 1.

1H NMR spectroscopy indicated successful synthesis of {3}, yielding chemical shifts characteristic of aromatic hydrogens as three doublets at 7.3–7.8 ppm, whereas the glucose CH protons resonated as six distinct multiplets at 3.9–6.0 ppm and the methyl groups of the protecting isopropylidenes resonated as four singlets at 1.2–1.6 ppm (Fig. S2).

Synthesis of tetra-β-glycosylated phthalocyanines

4-(1,2:3,4-Di-O-isopropylidene-α-d-glucopyranose)-phthalonitrile {3} then underwent cyclization in the presence of n-pentanol, Zn(OAc)2.2H2O and DBU, forming tetra-β-glycosylated ZnPc {4} (Scheme 1, step 2). The synthesis of tetrasubstituted phthalocyanines is known to produce mixtures of regioisomers (32) due to the various possible positions of the substituents, relative to one another (33). Thus, ZnPc {4} was presumably a mixture of regioisomers, with the glucose moiety on either of the two possible beta positions of each phthalonitrile aromatic ring, but no attempt was made to separate them, and the product, a blue–green amorphous solid was purified by silica gel column chromatography using ethyl acetate–n-hexane as the eluent.

Characterization of protected tetra-β-glycosylated ZnPc {4}

The 1H NMR spectrum of ZnPc {4} gave characteristic chemical shifts of aromatic protons of the phthalocyanine ring (34) as three multiplets at 7.6–9.3 ppm, whereas glucose protons resonated as six multiplets at 4.1–5.8 ppm and the protecting isopropylidene methyl groups resonated as four singlets at 1–2 ppm (Fig. S3A).

Characterization of deprotected tetra-β-glycosylated ZnPc {5}

The 1H NMR spectrum of ZnPc {5} (Fig. S4) also gave the characteristic chemical shifts of the aromatic phthalocyanine protons as three multiplets at 7.6–9.6 ppm. The glucose protons resonated at 4.0–5.5 ppm, but due to the broadening of the water peak in DMSO-d6, it was not possible to establish the exact number of glucose multiplets. The 1H NMR spectrum demonstrated that the deprotection of the isopropylidene methyl groups was largely successful as evidenced by the disappearance of CH3 groups at 1–2 ppm (compare with ZnPc {4} in Fig. S3A).

The calculated molecular weight of ZnPc {5} (C56H56N8O24Zn) was 1290.47, but the MALDI TOF MS gave a major signal at m/z 1466.2, which did not match the expected molecular ion (Fig. S5). The m/z 1466.2 signal is instead consistent with a partially deprotected ZnPc {5} containing (on average) four isopropylidene groups still attached (i.e.∼50% deprotection). The expanded MALDI TOF MS spectrum revealed the presence of the various mono (Fig. S6A) and dimeric (Fig. S6B) species of ZnPc {5}.

UV-Vis absorption spectroscopic properties

The electronic absorption of ZnPc {4} and {5} were recorded at increasing concentrations (1, 3, 6, 9 and 12 μm) in DMSO or phenol red-free DMEM (Fig. 1). In DMSO, the UV-Vis spectra of ZnPc {4} showed a B band at 357 nm, a vibronic band at 613 nm and a Q-band at 682 nm (Fig. 1A). ZnPc {5} also showed the three bands: a B band at 357 nm, a vibronic band at 617 nm and a Q band at 686 nm (Fig. 1B). The intense, single and narrow Q band is thought to be typical of monomeric metallated phthalocyanine complexes (33,35).

Figure 1. UV-Vis absorption spectra of (A, C) tetra-β-glycosylated ZnPc {4} and (B, D) ZnPc {5} showing the absence of aggregation at various concentrations in DMSO (A,B), but showing aggregation with increasing concentrations in DMEM (C, D). The inset graphs in each panel show the absorbance of different concentrations (1, 3, 6, 9 and 12 μm) of each compound in the same solvent at a fixed wavelength. (E, F) Fluorescence spectra of (E) ZnPc {4} and (F) ZnPc {5} at different concentrations (1, 3, 6, 9 and 12 μm) in DMSO. Em 365 nm. (G) Fluorescence spectra of 9 μm ZnPc {5} in water, in the presence or absence of 5% SDS detergent. Inset shows the expanded spectrum of ZnPc {5} in water. Em 365 nm..

Regardless of concentration, the Q band remained single and narrow for both compounds, an indication that no significant aggregation took place in DMSO (33). Moreover, when the maximum Q band absorbance (682 nm for ZnPc {4} or 686 nm for ZnPc {5}) was plotted against the different concentrations tested, straight line graphs were obtained (insets in Figs. 1A and B), obeying the Beer–Lambert law and further supporting the existence of ZnPc {4} and ZnPc {5} as monomers in DMSO (36).

In contrast to the well-defined absorbance spectra obtained in DMSO, the UV-Vis spectra of ZnPc {4} (Fig. 1C) and {5} (Fig. 1D) in an aqueous solution (DMEM) showed a broadened Q band with much lower absorbance compared with the same concentrations in DMSO. ZnPc {4} had two peaks with maximum absorbance at 637 and 678 nm (Fig. 1C) whereas ZnPc {5} had only one peak with maximum absorbance at 637 nm (Fig. 1D). The broadened and low-intensity Q bands suggested that ZnPc {4} (Fig. 1C) and {5} (Fig. 1D) were aggregated in DMEM media, as had been observed previously by other groups (37,38). However, the absorbance values for different concentrations (1, 3, 6, 9 and 12 μm), at fixed wavelengths, 674 nm for ZnPc {4} (Fig. 1C, inset) and at 637 nm for ZnPc {5} (Fig. 1D, inset), gave straight line graphs obeying the Beer–Lambert law, suggesting that a proportion of each compound remained as monomers in DMEM, in agreement with Lo et al. (16).

Fluorescence spectroscopic properties

Fluorescence spectra of ZnPc {4} and {5} were recorded at increasing concentrations (1, 3, 6, 9 and 12 μm) in DMSO (Fig. 1E,F) or phenol red-free DMEM (not shown). In DMSO, ZnPc {4} and {5} showed fluorescence intensity maxima at 700 and 704 nm, respectively. However, in DMEM both compounds exhibited negligible fluorescence (not shown). The poor fluorescence of these compounds in a polar solution, such as DMEM, is likely a result of aggregation, leading to fluorescence quenching (39,40). In support of this hypothesis, strong phthalocyanine fluorescence was restored when 5% SDS detergent was added to 9 μm ZnPc {5} in water (Fig. 1G). To better understand the possible molecular mechanisms of aggregation, we used RLS. RLS data demonstrated moderate concentration-dependent electronic coupling between adjacent ZnPc {4} molecules, sufficient to initiate limited aggregation in water-based solutions at concentrations above about 6 μm that produced an RLS signal (Fig. 2). No RLS signal was detected in solutions of ZnPc {5} (not shown).

Figure 2. RLS signals of different concentrations of ZnPc {4} in (A) water, (B) PBS and (C) DMEM. (D) RLS signals from the 5 μm MC540 aggregation positive control in each of the aqueous media.

The extent of ZnPc {4} aggregation was similar in all the aqueous media tested, but the RLS signal was always less pronounced than the aggregation positive control of 5 μm MC540. However, no quantitative comparisons of the size or quantity of aggregates can be made between these compounds using RLS. Thus, ZnPc {4} aggregates can be detected in DMEM, even when Beer–Lambert law plots are almost linear. Taken together, the fluorescence and absorbance data indicate that ZnPc {4} and {5} are aggregated to some extent in DMEM, although the type of aggregation may differ. Thus, ZnPc {5}, which does not produce an RLS signal, must be aggregated via nonelectronic coupled forces, such as hydrogen bonding through exposed –OH (hydroxyl) groups on the glucose moieties and/or hydrophobic interactions.

Cellular uptake and accumulation of glycosylated PSs

Having successfully synthesized and chemically characterized ZnPc {4} and having established that ZnPc {4} and {5} were glycoconjugated phthalocyanines, the next steps were to assess their in vitro uptake kinetics, subcellular localization, dark toxicity, phototoxicity and ROS generation in MCF-7 cells, in comparison with the well-characterized second-generation PS AlPcS2 (41). In addition, we assessed whether the aggregation displayed by our compounds in DMEM impacted negatively on their photodynamic activity in vitro,.

Because ZnPc {4} and {5} are glycosylated, we predicted that they would preferentially be taken into MCF-7 cells by facilitated diffusion via GLUT receptors, in contrast to the nonglycosylated AlPcS2. If this was the case, then the uptake of ZnPc {4} and {5}, but not of AlPcS2, should be competitively inhibited in the presence of excess (25 mm) glucose, compared with incubation in physiologically relevant 5 mm glucose. However, although the uptake and accumulation of ZnPc {4} was up to 10-fold higher than that of equimolar AlPcS2, there was no significant inhibition of uptake in 25 mm glucose (Fig. 3A), suggesting that GLUT receptors are not the primary method of uptake and accumulation for ZnPc {4}. Indeed, our data indicated that, between 2 and 4 h of incubation, the uptake of ZnPc {4} became greater in 25 mm glucose, compared with 5 mm glucose. This was not a general feature, as the uptake of AlPcS2 was slightly lower in 25 mm glucose compared with 5 mm glucose throughout the period of incubation (Fig. 3B). By contrast, ZnPc {5} showed negligible levels of uptake (maximum uptake 0.075 pm per cell; 100-fold less than ZnPc {4}). Acidification of the cytoplasm or culture medium, due to lactic acid generation as a consequence of increased glycolysis (42), might account for the increased uptake of ZnPc {4} in the presence of 25 mm glucose. Previously, it was shown that a decrease in pH increased the cellular uptake of lipophilic PSs (43,44).

Figure 3. Cellular uptake of ZnPc {4} (red), ZnPc {5} (green) and AlPcS2 (blue) as a function of incubation time. MCF-7 cells were incubated with 3 or 9 μm of the specified photosensitizers for up to 4 h in 5 mm glucose (continuous lines) or 25 mm glucose (dashed lines). Figure (A) contains the plots of all the photosensitizers, whereas (B) shows an expanded plot for AlPcS2 and ZnPc {5}. Each point represents the mean of three independent experiments (mean ± SEM), each one performed in triplicate.

Cytotoxicity studies

The effects of ZnPc {4} and {5} on MCF-7 cell viability and cell number in the absence of light (dark toxicity) and after PDT treatment (phototoxicity) were investigated and compared with AlPcS2. In these cytotoxicity studies, cells were loaded for 2 h with 0, 1, 3, 6 or 9 μm of either ZnPc {4} or AlPcS2 and analyzed 24 h later for cell morphology, PI exclusion assay and total cell number. Due to the poor uptake and accumulation of ZnPc {5}, only the highest concentration of this PS (9 μm) was included in the study.

Morphology. The phase contrast micrographs showed that for all the concentrations investigated, ZnPc {4} did not produce any obvious dark toxicity, as there was no increased cell detachment or change in morphology of the MCF-7 cells (Fig. 5A). By contrast, at the highest concentrations (6 and 9 μm), AlPcS2 produced some detachment of cells in the dark (Fig. 5B). Twenty-four hours following PDT, both ZnPc {4} (Fig. 5A) and AlPcS2 (Fig. 5B) showed a concentration-dependent increase in floating cells and an increase in shrunken and apparently ruptured cells. AlPcS2 treatment showed floating cells at concentrations of 3 μm and above, whereas ZnPc {4} treatment showed floating cells at concentrations of 6 μm and above. By contrast, 9 μm ZnPc {5} demonstrated a slight increase in the number of floating cells, but no obvious cell morphology changes in the attached cells, compared with PS-free controls (Fig. 6). However, the morphology of the attached cells was not a faithful predictor of their viability. Thus, using ZnPc {4} concentrations of 3 μm and above or AlPcS2 concentrations of 6 μm and above, 24 h after PDT, we detected several attached cells with an apparently normal morphology that were permeable to PI and conversely several shriveled cells that remained impermeable to PI (Fig. 5C). It is possible that the PI-permeable cells had been quickly killed by PDT treatment, without time to undergo morphological changes or to float off, analogous to histological fixation.

Figure 5. MCF-7 cells were treated with different concentrations (0, 1, 3, 6, 9 μm) of (A) ZnPc {4} or (B) AlPcS2 for 2 h before exposure to light for 15 min (or kept in the dark) and were then photographed (20× objective) after 24 h. (C) Some light-exposed cells were stained with propidium iodide (PI) and merged brightfield/PI fluorescence (red) images are shown. Note that cell morphology is not a good predictor of cell viability under these conditions. Live cells are indicated by a white arrowhead and adjacent dead (PI+) cells with a similar morphology are identified by a black arrowhead.

Figure 6. MCF-7 cells were treated with 0 μm (control) or 9 μm ZnPc {5} for 2 h before exposure to light for 15 min (or kept in the dark). The representative phase contrast images (20× objective) were a snapshot of the cells before analysis of the cell viability by PI exclusion assay and cell count.

Cell viability. Cell viability was quantified by PI exclusion assay and the results were expressed as a percentage of the total cell population (Fig. 7A,B). In the dark, the treatment of MCF-7 cells with 1, 3, 6 or 9 μm ZnPc {4} or with 9 μm ZnPc {5} did not significantly affect cell viability over the 24 h duration, compared with the dark no PS control (Fig. 7A). AlPcS2 did not significantly affect cell viability in the dark at either 1, 3 or 6 μm, but it did significantly (P <0.01) decrease cell viability at 9 μm, compared with no PS controls (Fig. 7A).

Figure 7. Dark toxicity and phototoxicity effects of photosensitizers on (A,B) cell viability, (C,D) total cell number and (E) viability ratio. MCF-7 cells were incubated for 2 h with the specified concentrations of ZnPc {4} (red), ZnPc {5} (green) or AlPcS2 (blue), then exposed to light for 15 min (or kept in the dark for the dark toxicity studies). Twenty-four hours later, a PI exclusion FACS assay was carried out to determine the cell viability and the total cell number per well was determined by hemocytometer. (C) Viability ratios were calculated from cell survival and cell number data from the dark and phototoxicity studies. Dotted line (ratio of 1) indicates no difference in cell viability between the dark and phototoxicity. In each case, results represent the mean of at least three independent experiments per condition (mean ± SEM) and were analyzed by two-way repeated-measures ANOVA with Bonferroni posttests for cell viability and viability ratio data, and by one-way ANOVA with Dunnett’s multiple comparison test for total cell numbers. The symbols denote statistically significant differences compared with control (P <0.05*, P <0.01**, P <0.001***) or (in A,B,E) compared with ZnPc {4} (P <0.001###).

The incubation of MCF-7 cells with either 1 or 3 μm ZnPc {4}, followed by PDT, slightly, but nonsignificantly, decreased cell viability by 24 h compared with light-exposed no PS control samples, with over 90% of the cell populations remaining PI negative (Fig. 7B). By contrast, incubation with 6 or 9 μm ZnPc {4} significantly (P <0.001) decreased cell viability compared with light-exposed no PS control samples with 9 μm ZnPc {4} decreasing the cell viability by more than 50% (Fig. 7B). AlPcS2 followed the same pattern as ZnPc {4}, with 1 and 3 μm AlPcS2 hardly affecting the cell viability, whereas 6 or 9 μm AlPcS2 significantly (P <0.001) decreased cell viability compared with light-exposed no PS control samples (Fig. 7B). Importantly, 6 or 9 μm ZnPc {4} caused significantly (P <0.001) more PDT cell death when compared with equimolar 6 or 9 μm AlPcS2. The trends in cell death from PI/FACS (Fig. 7) are consistent with our PI/brightfield data (Fig. 5C).

Total cell numbers. In the dark, none of the PSs, at any concentration, decreased total cell number per well by more than 5% compared with no PS control (Fig. 7C). The incubation of MCF-7 cells with increasing concentration of ZnPc {4} (1, 3, 6 and 9 μm), followed by PDT, decreased the cell number per well by 0.3%, 13.0%, 45.2% (P <0.01) and 57% (P <0.01), respectively, compared with light-exposed no PS control samples (Fig. 7D). AlPcS2 followed the same pattern, with increasing concentration (1, 3, 6 and 9 μm) decreasing the cell number per well by 4.2%, 19.7%, 34.9% (P <0.01) and 44.0% (P <0.01), respectively, compared with light-exposed no PS control samples (Fig. 7D). Cells incubated with 9 μm ZnPc {5} showed an 8.8% reduction in cell number per well following PDT, which was not significant compared with light-exposed no PS control samples (Fig. 7D).

PDT-specific cell death (viability ratios). To accurately quantify PDT-specific cell death for each PS, it was necessary to correct for any inherent (dark) toxicity. To achieve this, we calculated the total number of viable cells per well from their respective percentage cell viability (Fig. 7A,B) and total cell number per well data (Fig. 7C,D). By dividing the paired values for total viable cells following PDT treatment by total viable cells in the dark, a viability ratio was obtained (Fig. 7E).

A dose-dependent decrease in viability ratio was observed for ZnPc {4} and AlPcS2 and these decreases became significant compared with no PS controls at 3 μm and above (Fig. 7E). Thus, incubation with 3, 6 or 9 μm ZnPc {4} lowered the viability ratios to 0.83, 0.41 and 0.22, respectively, indicating that PDT decreased the number of viable cells by 17%, 59% and 78%, respectively, compared with paired samples in the dark. Similarly, incubation with 3, 6 or 9 μm AlPcS2 decreased the viability ratios to 0.75, 0.60 and 0.47, respectively, indicating that PDT decreased the number of viable cells by 25%, 40% and 53%, respectively, compared with paired samples in the dark. Importantly, ZnPc {4} demonstrated significantly lower viability ratios than AlPcS2 at 6 μm (P <0.001) and 9 μm (P <0.001), with ZnPc {4} showing a 25% increase in cell death over AlPcS2 at 9 μm (Fig. 7E).

Intracellular ROS levels

Intracellular ROS levels were assayed by the oxidation of DCFH-DA to fluorescent DCF. The results were normalized to the light-exposed no PS sample (light control) by dividing the mean DCF fluorescence of each test sample by the mean DCF fluorescence of the light control sample.

All test samples maintained in the dark showed ROS levels on average 90% lower (Fig. 8A) than the light control level (Fig. 8B), with no significant differences between them, suggesting no intrinsic differences in DCFH-DA uptake and accumulation in the presence of the various PSs.

Incubation with 9 μm AlPcS2, followed by light exposure, significantly increased (P <0.01) the general ROS levels by an average of 229.6%, compared with the light control (Fig. 8B). Very encouragingly, incubation with 9 μm ZnPc {4}, followed by light exposure, significantly increased (P <0.001) the general ROS levels by 758.1% compared with light control and by 528.5% compared with light-treated AlPcS2 samples (P <0.001) (Fig. 8B). These ROS results are consistent with the higher uptake and accumulation of ZnPc {4}, compared with AlPcS2 and likely explain the improved phototoxicity of ZnPc {4} on MCF-7 cells. By contrast, incubation with 9 μm ZnPc {5} followed by light exposure only increased the general ROS levels by an average of 9.9%, which was not significantly different to the light control (Fig. 8B).

Discussion

Glycosylated phthalocyanines have attracted interest recently (1,14,17,35,45–47). The rationale being that the carbohydrate moiety (e.g. glucose) should increase solubility and also target the PSs to GLUT receptors, generally up-regulated on tumor cells, thus solving the persistent problems regarding the poor cellular uptake, low tumor selectivity and lower than hoped for photosensitizing ability of previous phthalocyanine-based PSs (20). The aim of this study was to synthesize and characterize a tetra-β-glycosylated Zinc (II) phthalocyanine. Although we were successful in generating a partially deprotected version of the desired compound (ZnPc {5}), we found, unexpectedly, that the intermediate isopropylidene-protected compound (ZnPc {4}) had far superior uptake, PDT–ROS generation and PDT cytotoxicity in MCF-7 cells than either ZnPc {5} or AlPcS2. Using equimolar (9 μm) concentrations of each compound and standardized illumination conditions (28.6 J cm−2), it was evident that ZnPc {4} was the most phototoxic to MCF-7 cells (9 μm giving: LD75 based on viability ratio, and IC50 based on percentage viability), followed by AlPcS2 (LD50 9 μm), whereas ZnPc {5} showed much lower phototoxicity (LD10 9 μm). This paralleled our observed differences in cellular uptake and PDT ROS generation and is consistent with other studies showing a relationship between PS uptake and PDT cell kill (15,21,45). We have not measured parameters such as singlet oxygen quantum yield, but the simplest explanation for the superior PDT cytotoxicity of ZnPc {4} is greater light absorption and consequent ROS generation, due to its high cellular uptake. Despite the presence of glycosylated substituents on the phthalocyanine macrocycle, the uptake of ZnPc {4} and {5} by MCF-7 cells was not primarily via GLUTs, as the uptake was not competitively inhibited by excess glucose. Uptake may have been promoted by zinc chelation, which has been shown to increase membrane-binding efficiency of porphyrin PSs (48). We noticed large amounts of ZnPc {4} attached to cell surfaces during incubation, both by phase illumination (not shown) and by confocal microscopy (Fig. 4). However, this does not explain why ZnPc {5} is so poorly taken up by cells, unless it is aggregated in such a way that sterically blocks any beneficial effect of zinc chelation.

A previous study, using a similar synthetic route to make tetra-β-glycosylated ZnPc, found negligible cellular uptake of the protected form (1), in stark contrast to the excellent uptake and phototoxicity we found for protected ZnPc {4}. Specifically, Choi et al. (1) found that protected tetra β-glycosylated ZnPcs were photochemically inactive, whereas protected mono-β-glycosylated ZnPc had PDT IC50 values between 0.9 and 1.5 μm in human adenocarcinoma and hepatoma cell lines, using a high light dose (48 J cm−2). The main difference between their synthetic route and ours was that we used a higher reaction temperature (120°C) for each step (see Scheme 1), whereas the previous study used 60°C or room temperature during the various steps (1). This likely resulted in us producing a different and perhaps more varied mixture of ZnPc {4} regioisomers than had been made previously and it will be important in future studies to determine which isomers are preferentially taken up by MCF-7 cells in order to direct future rational PS design.

Surprisingly, our deprotected compound ZnPc {5} demonstrated negligible cellular uptake, even though it was more water soluble than ZnPc {4}. This may have been due to more stable aggregation of ZnPc {5}. ZnPc {5} did not produce any RLS signal, indicating that aggregation was not due to electronic coupling. However, the addition of 5% SDS to an aqueous solution of ZnPc {5} restored strong fluorescence, confirming aggregation-induced fluorescence quenching. Hydrogen bonding alone is unlikely to account for the aggregation in water, because the OH groups on the glucose would be solvated by water molecules through H-bonding. More likely, aggregation of ZnPc {5} is driven by hydrophobic stacking between the aromatic rings of phthalocyanine, allowing the free OH groups on glucose to form H-bonds with each other as nearest neighbors, further stabilizing the aggregates, as is seen in similar phthalocyanine compounds (49). Liu et al. (35) similarly reported that a large number of substituents at the β-positions exaggerates phthalocyanine aggregation and, in agreement with our work, that removal of isopropylidene-protecting groups reduces phototoxicity. Our initial and encouraging data now pave the way for further studies to better understand the mechanism of ZnPc {4} uptake, to overcome solubility issues by improving the formulation, and to move our work into a wider variety of tumor models.

Liu, J. Y., P. C. Lo, W. P. Fong and D. K. Ng (2009) Effects of the number and position of the substituents on the in vitro photodynamic activities of glucosylated zinc(II) phthalocyanines. Org. Biomol. Chem.7, 1583–1591.

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